Scholarly article on topic 'Intrinsic Membrane Hyperexcitability of Amyotrophic Lateral Sclerosis Patient-Derived Motor Neurons'

Intrinsic Membrane Hyperexcitability of Amyotrophic Lateral Sclerosis Patient-Derived Motor Neurons Academic research paper on "Biological sciences"

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Abstract of research paper on Biological sciences, author of scientific article — Brian J. Wainger, Evangelos Kiskinis, Cassidy Mellin, Ole Wiskow, Steve S.W. Han, et al.

Summary Amyotrophic lateral sclerosis (ALS) is a fatal neurodegenerative disease of the motor nervous system. We show using multielectrode array and patch-clamp recordings that hyperexcitability detected by clinical neurophysiological studies of ALS patients is recapitulated in induced pluripotent stem cell-derived motor neurons from ALS patients harboring superoxide dismutase 1 (SOD1), C9orf72, and fused-in-sarcoma mutations. Motor neurons produced from a genetically corrected but otherwise isogenic SOD1 +/+ stem cell line do not display the hyperexcitability phenotype. SOD1 A4V/+ ALS patient-derived motor neurons have reduced delayed-rectifier potassium current amplitudes relative to control-derived motor neurons, a deficit that may underlie their hyperexcitability. The Kv7 channel activator retigabine both blocks the hyperexcitability and improves motor neuron survival in vitro when tested in SOD1 mutant ALS cases. Therefore, electrophysiological characterization of human stem cell-derived neurons can reveal disease-related mechanisms and identify therapeutic candidates.

Academic research paper on topic "Intrinsic Membrane Hyperexcitability of Amyotrophic Lateral Sclerosis Patient-Derived Motor Neurons"

Cell Reports


Intrinsic Membrane Hyperexcitability of Amyotrophic Lateral Sclerosis Patient-Derived Motor Neurons

Brian J. Wainger,128 Evangelos Kiskinis,38 Cassidy Mellin,1 Ole Wiskow,3 Steve S.W. Han,34 Jackson Sandoe,3 Numa P. Perez,1 Luis A. Williams,3 Seungkyu Lee,1 Gabriella Boulting,3 James D. Berry,4 Robert H. Brown, Jr.,5 Merit E. Cudkowicz,4 Bruce P. Bean,6 Kevin Eggan,3 4 7 * and Clifford J. Woolf16 *

1FM Kirby Neurobiology Center, Boston Children's Hospital and Harvard Stem Cell Institute, Boston, MA 02115, USA

2Department of Anesthesia, Critical Care and Pain Medicine, Massachusetts General Hospital, Boston, MA 02114, USA

3Harvard Stem Cell Institute, Department of Stem Cell and Regenerative Biology, Harvard University and the Stanley Center for Psychiatric

Research, Broad Institute, Cambridge, MA 02138, USA

4Department of Neurology, Massachusetts General Hospital, Boston, MA 02114, USA 5Department of Neurology, University of Massachusetts Medical Center, Worcester, MA 01655, USA 6Department of Neurobiology, Harvard Medical School, Boston, MA 02115, USA 7The Howard Hughes Medical Institute 8These authors contributed equally to this work

'Correspondence: (K.E.), (C.J.W.)

This is an open access article under the CC BY-NC-ND license (


Amyotrophic lateral sclerosis (ALS) is a fatal neurodegenerative disease of the motor nervous system. We show using multielectrode array and patch-clamp recordings that hyperexcitability detected by clinical neurophysiological studies of ALS patients is recapitulated in induced pluripotent stem cell-derived motor neurons from ALS patients harboring superoxide dismutase 1 (SOD1), C9orf72, and fused-in-sarcoma mutations. Motor neurons produced from a genetically corrected but otherwise isogenic SOD1+'+ stem cell line do not display the hyperexcitability phenotype. SOD1A4V/+ ALS patient-derived motor neurons have reduced delayed-rectifier potassium current amplitudes relative to control-derived motor neurons, a deficit that may underlie their hyperexcitability. The Kv7 channel activator retigabine both blocks the hyperexcitability and improves motor neuron survival in vitro when tested in SOD1 mutant ALS cases. Therefore, electrophysiological characterization of human stem cell-derived neurons can reveal disease-related mechanisms and identify therapeutic candidates.


Amyotrophic lateral sclerosis (ALS) is a devastating, untreatable disease of upper and lower motor neurons (Kiernan et al., 2011). The excitotoxicity neurodegeneration hypothesis posits that excessive glutamatergic synaptic activity in ALS leads to calcium overload and cell death (Cleveland and Rothstein, 2001; Pasinelli and Brown, 2006). However, nerve conduction studies

evaluating axonal threshold (strength-duration time constant and recovery cycle times) in ALS patients demonstrate increased axonal membrane excitability, well away from any synapses (Bostock et al., 1995; Kanai et al., 2006; Nakata et al., 2006; Vucic and Kiernan, 2006), and the degree of hyperexcitability correlates with patient survival (Kanai et al., 2012). Increased membrane excitability may be important then as a contributor to disease, and modeling suggests that either increased persistent sodium or reduced delayed-rectifier potassium currents could be responsible for the axonal hyperexcitability (Kanai et al., 2006; Tamura et al., 2006). However, whether excitability results from autonomous changes in motor neurons cannot be determined by this technique (Fritz et al., 2013).

About 10% of ALS cases are familial, and, of these, superoxide dismutase 1 (SOD1) mutations account for about 20%. Motor neurons from SOD1G93A mice, which overexpress this human mutant SOD1 protein, also show hyperexcitability (Kuo et al., 2004; Pieri et al., 2003; van Zundert et al., 2008), at least in part due to increased persistent sodium currents. Because of the distinct clinical and pathological features of SOD1 ALS compared to other variants (Ince et al., 2011), it is unclear if primary motor neuron hyperexcitability represents a general feature of ALS or a specific characteristic of SOD1 -mediated disease. Hyperexcitability in motor neurons of other familial ALS etiologies, such as C9orf72 hexanucleotide repeat expansions and fused-in-sarcoma (FUS) mutations, have not yet been similarly evaluated because of a lack of mouse models.

Induced pluripotent stem cell (iPSC) technology enables neurons of specific disease-relevant subtypes to be derived from disease patients and control subjects and thereby provides an in vitro platform for discovering human neuron phenotypes that may reflect the individual diseases. However, the number of subject cell lines employed in studies utilizing this technique has so far been small (Sandoe and Eggan, 2013). Thus, it is currently difficult to know how consistent such findings are across large

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numbers of patients, and if the results represent disease-specific phenotypes or differences among cell lines. We address these issues here in two ways. First, we use a gene-targeted correction of the disease-causing SOD1 mutation to produce otherwise isogenic stem cells bearing wild-type SOD1 alleles (Kiskinis et al., 2014) and show that the gene correction abrogates the phenotype of ALS motor neuron hyperexcitability. Second, we demonstrate that hyperexcitability is present among motor neurons derived from eight ALS patients of three separate genetic etiologies compared to five control patients, together constituting the largest sample group to date for these types of studies. ALS-derived motor neurons have reduced delayed-rectifier voltage-gated potassium currents compared to controls. Furthermore, retigabine, an activatorof Kv7 potassium channels reduces excitability to levels seen in controls. Retigabine increases the in vitro survival of SOD1A4V/+ ALS motor neurons, supporting the hypotheses that motor neuron hyperexcitability may contribute to motor neuron degeneration in ALS. The hyperexcitability phenotype and blockade of firing by retigabine are present across a wide range of familial ALS patients harboring additional SOD1 mutations, C9orf72 repeat expansions, and FUS mutations. An iPSC-based disease-modeling approach can validate a clinically relevant phenotype in human motor neurons, reveal mechanisms underlying the phenotype, and help evaluate actions of candidate drugs on the disease-specific phenotype.


Hyperexcitability of SODiA4V-Derived Motor Neurons Using Multielectrode Array Recording

We performed an initial set of electrophysiological phenotyping experiments using iPSC-derived motor neurons from two control subjects (11a, 18a) and two unrelated familial ALS patients (39b and RB9d) harboring the same aggressive SOD1A4V/+ mutation. All iPSC lines were generated via three-factor (OCT4, SOX2, KLF4) retroviral reprogramming, had a normal karyotype, and differentiated into motor neurons after robust neuralization based on dual SMAD inhibition (Chambers et al., 2009) and specification through exposure to retinoic acid and induction of sonic hedgehog signaling (Boulting et al., 2011; Kiskinis et al., 2014; Figure 1A).

We recorded spontaneous firing of iPSC-derived motor neurons using extracellular multielectrode arrays (MEAs), whereby the action potentials of individual neurons are detected by a grid of 64 extracellular electrodes embedded in each culture well. In four separate experiments, control and SOD1 ALS iPSC lines were cultured synchronously, differentiated into motor neurons in parallel, and plated in equal numbers on MEAs, which allowed recording of the spontaneous firing (Hanson and Landmesser, 2004) in hundreds of control and ALS patient-derived neurons per differentiation after culturing for 4 weeks. We observed significantly more spontaneous action potentials in SOD1A4V/+ relative to control cultures (p < 0.05, t test; Figures 1B and 1C). Action potentials were sorted by spike morphology and timing to derive clusters corresponding to individual neurons (Figure S1) (Cohen and Kohn, 2011), and a significantly higher average mean firing rate was observed in the SOD1A4V/+ neurons (p < 10~15, t test; Figures 1D and 1E).

We performed two experiments to confirm that the hyperexcitability on the MEAs resulted from motor neurons. First, to test if the difference in spontaneous action potential firing resulted from a larger number or more active population of inhibitory neurons in control cultures, we applied GABAergic and glycinergic transmission blockers. The blockers did not increase action potential firing rates (p = 0.61, t test, for bicuculline; p = 0.24, t test, for strychnine; Figure S2), suggesting that activity of inhibitory neurons was minimal and that the heightened spontaneous firing in the MEA recordings of ALS-derived motor neurons reflected an intrinsic increase in excitability.

Second, we inserted a Hb9::GFP reporter into the AAVS1 locus for iPSCs 18a and 39b (Figure S3A), allowing fluorescence-activated cell sorting (FACS) purification of GFP-positive motor neurons for recording on the arrays (Figures S3B and S3C). We recorded every 4 days and observed that 39b Hb9::GFP motor neurons consistently fired more action potentials than 18a Hb9::GFP motor neurons over the entire time course (mixed model ANOVA F-test p = 1 x 10~4 for difference between lines; post hoc t tests after Bonferroni correction for multiple comparisons for day 12, p = 0.0055; day 16, p = 0.0030; day 20, p = 0.0057; day 24, p = 0.0029; day 28, p = 0.015; Figure 1F). Thus, the hyperexcitability must be due to motor neurons, because only Hb9-positive motor neurons were plated on the arrays.

Correction of the SOD1A4V Mutation Eliminates the Hyperexcitability Phenotype

To test if increased action potential firing was a direct effect of the SOD1A4V mutation, we took advantage of a gene-targeted derivative of the 39b iPSC line in which the A4V-encoding mutation had been corrected to a wild-type sequence by homologous recombination, 39b-SOD1+/+ (abbreviated 39b-Cor; Kiskinis et al., 2014). Because substantial motor neuron death begins in ALS motor neurons after 15 days of neuronal maturation in our culture conditions (Kiskinis et al., 2014) and because the hy-perexcitability phenotype in the ALS motor neurons was detectable at this early time point (Figure 1F), we compared MEA recordings of 39b and isogenic-derived 39b-Cor motor neurons at 14 days, to avoid the possibility that increased firing reflected either neuronal death or select survival of hyperexcitable neurons (Table S1). Although the baseline spike rate was lower at 14 than 28 days, patient-derived 39b neurons had a higher spontaneous firing rate than neurons in which the SOD1 mutation was corrected (p = 0.01 for total rate, t test; Figure 1G; average mean firing rate 1.30 ± 0.10 Hz for n = 122 39b-Cor and 1.50 ± 0.08 Hz for n = 208 39b; p < 0.05, t test). We conclude that the hyperex-citability phenotype reflects the presence of the disease-initiating mutation and precedes progressive motor neuron death.

Confirmation of ALS Motor Neuron Hyperexcitability and Mechanistic Exploration Using Whole-Cell Patch Clamp

To examine the electrophysiological properties of identified individual motor neurons, we transduced developing neurons with an Hb9::RFP lentiviral reporter and recorded only from RFP-pos-itive motor neurons (Marchetto et al., 2008) using whole-cell patch clamp (Figure 2A). Both control- and SOD1 ALS-derived motor neurons were electrically excitable. To quantify the degree

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Figure 1. Multielectrode Array Recording Reveals Increased Spontaneous Firing in ALS-Derived Neurons Compared to Control-Derived Neurons

(A) Schematic of differentiation and recording.

(B) Representative recordings from four out of 64 MEA electrodes in control (11a, 18a) and ALS (39b, RB9d)-derived neurons cultured for 28 days on the arrays.

(C) Total action potential firing rate during 1 min of recording from MEAs (11a, n = 3; 18a, n = 3; control mean 6,510 ± 3,131 spikes/min; 39b, n = 3; RB9d, n = 3; ALS mean 20,528 ± 5,069 spikes/min; p < 0.05, t test).

(D) Mean firing rate histograms of individual neurons from MEAs in (B). See also Figure S1.

(E) Average of mean firing rate for patient-derived neurons (11a, n = 381; 18a, n = 191; control mean 1.17 ± 0.04 Hz; 39b, n = 520; RB9d, n = 662; ALS mean 1.76 ± 0.05 Hz; p< 10~15, t test).

(F) Total action potential firing rate during 1 min recordings from MEAs of FACS-sorted 18a Hb9::GFP and 39b Hb9::GFP motor neurons recorded every 4 days (repeated-measures ANOVA F-test p = 1 x 10~4 for difference between lines; post hoc t tests with Bonferroni correction for multiple testing indicated as *p < 0.05 and **p < 0.01). See also Figures S2 and S3.

(G) Total action potential firing rate during 1 min of recording from MEAs cultured for 14 days on the arrays (39b-Cor, n = 4; mean 775 ± 712 spikes/min; 39b, n = 4; 39b mean 6,278 ± 1,758 spikes/min; p = 0.01, t test).

Error bars represent SEM.

of excitability, we assayed the number of action potentials fired in response to a slow ramp depolarization. The number of action potentials fired by ALS motor neurons was significantly greater than control motor neurons (p < 0.05, Mann-Whitney U test; Figure 2B, upper panels, Figure 2C, upper panel). Resting membrane potential, action potential threshold (rheobase), and input resistance did not differ between ALS and control motor neurons (Table S2), indicating that excitability differences were not due to differences in electrophysiological health or baseline capacity for action potential generation.

When we compared motor neurons derived from 39b-Cor and 39b cell lines in three separate additional parallel experiments,

we again observed a marked difference in the number of action potentials elicited during ramp depolarization (p < 0.05, MannWhitney U test; Figure 2B, lower panels, Figure 2C, lower panel), demonstrating that the A4V mutation was essential for the phenotype. There was variability in the number of action potentials in motor neurons from the same line tested across multiple differentiations, but the increased number of action potentials in ALS motor neurons relative to control motor neurons was always preserved. This result underscores the importance of performing repeated parallel differentiations in which equal numbers of control and ALS motor neurons are analyzed from each differentiation.

Figure 2. ALS Patient-Derived Motor Neurons Are Hyperexcitable and Have Reduced Delayed-Rectifier Potassium Currents Compared to Control-Derived Motor Neurons

(A) An iPSC-derived motor neuron identified by Hb9::RFP lentiviral transduction (right) and during patch-clamp recording (left) after culture for 28 days. Scale bar, 20 mm.

(B) Representative current clamp recordings during ramp depolarization from control and ALS patient-derived motor neurons (upper four panels); sample recordings from separate experiments comparing the isogenic correction of the 39b SOD1A4V mutation (39b-Cor) and 39b (lower two panels).

(C) Upper panel: average number of action potentials elicited by ramp depolarization from control (11a, n = 12; 18a, n = 11; control mean 2.5 ± 0.4) and ALS (39b, n = 13; RB9d, n = 12; ALS mean 4.2 ± 0.5) motor neurons obtained from four separate differentiations (p < 0.05, Mann-Whitney U test). Lower panel: separate experiments showing average number of action potentials during ramp depolarization from 39b-Cor (n = 17; mean 4.1 ± 0.5) and 39b (n = 19; mean 6.4 ± 0.9) motor neurons from three additional differentiations (p < 0.05, Mann-Whitney U test).

(D) Sample voltage-clamp recordings from control and ALS-derived Hb9::RFP-positive motor neurons cultured for 28 days.

(E) Average delayed-rectifier (DR) steady-state potassium current amplitude relative to peak sodium current amplitude in control (11a, n = 12; 18a, n = 11; control mean 0.88 ± 0.087) and ALS (39b, n = 13; RB9d, n = 12; ALS mean 0.44 ± 0.054) patient-derived motor neurons from four differentiations (p < 0.001, t test).

(F) Experiments from three separate differentiations showing average delayed-rectifier steady-state potassium current amplitude relative to peak sodium current amplitude in 39b-Cor (n = 18; mean 0.54 ± 0.061) and 39b (n = 19; mean 0.32 ± 0.036; p < 0.005, t test).

(G) Direct measurement of delayed-rectifier voltage-gated potassium current isolated by holding at —30 mV, stepping to a test-potential of +40 mV for 2 s and normalizing steady state current amplitude to cell capacitance in 39b-Cor (n = 19; mean 42.6 ± 4.3 pA/pF) and 39b (n = 18; mean 30.3 ± 3.1 pA/pF; p < 0.05, t test) derived motor neurons using cells from two additional separate differentiations.

(H) Peak sodium current amplitude normalized to cell capacitance in 39b-Cor (n = 16; mean 400.4 ± 44.7 pA/pF) and 39b (n = 15; mean 387.1 ± 50.5 pA/pF; p = 0.8, t test) derived motor neurons.

Error bars represent SEM.

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In addition to quantifying the electrical excitability of individual neurons, patch-clamp recording enables quantitative investigation of specific currents that determine excitability. To identify electrophysiological mechanisms responsible for the increased firing of mutant motor neurons, we performed voltage-clamp experiments using Hb9::RFP-positive motor neurons to examine current components. As an index of excitatory and inhibitory voltage-dependent ion channels, we quantified the ratio of outward delayed-rectifier potassium current to inward transient sodium current. In four repeated differentiations of motor neurons from control and SOD1A4V/+ iPSC lines, we observed that the ratio of delayed-rectifier potassium to transient sodium current was consistently smaller in SOD1A4V/+ motor neurons (p < 0.001, t test; Figures 2D and 2E). The difference was driven primarily by the reduced delayed-rectifier potassium channel component, as the difference in steady-state potassium current amplitude normalized to individual cell capacitance between ALS subjects and healthy controls was significant (control 137.0 ± 14.4 pA/pF, n = 23, versus ALS 94.4 ± 10.7 pA/pF, n = 25; p < 0.05, t test), whereas the peak sodium current normalized to capacitance was not (control 190.3 ± 23.0 pA/pF, n = 21, versus ALS 237 ± 21.2 pA/pF, n = 23; p = 0.2, t test). Because voltage-gated potassium channels repolarize the membrane

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(A) Rheobase measurements in a 39b Hb9::RFP-positive ALS-derived motor neuron in whole-cell patch clamp before (left) and after (right) the application of 10 mM retigabine (baseline rheobase 4.8 ± 1.5 pA versus post-retigabine rheobase 8.4 ± 2.2 pA; n = 11; p < 0.05, Wilcoxon signed rank test).

(B) Representative current clamp recording showing effect of 10 mM retigabine on membrane voltage and spontaneous firing (baseline Vm —60.4 ± 2.9 mV versus post-retigabine Vm —66.3 ± 3.6 mV, n = 11; p = 0.001, t test). In (A) and

(B), CNQX (15 mM), D-AP5 (20 mM), bicuculline (25 mM), and strychnine (2.5 mM) were added to the external solution.

(C) Dose-response curve for retigabine on suppression of spontaneous action potentials in MEA recording and Hill plot fit of mean data from 39b (n = 4) and RB9d (n = 4) with EC50 1.5 ± 0.8 mM.

(D) Effect of vehicle (open circles) and 1 mM retigabine (filled circles) treatment from days 14-28 of culture on the survival of Islet-positive, Tuj1-positive motor neurons measured at day 30 (total control n = 11; total ALS n = 9; F-test for effect of retigabine on all cells p = 3.8 x 10—4; effect of retigabine in ALS motor neurons, red, 25.3% (SD 5.6; t test p = 6.4 x 10—5); effect of retigabine in control motor neurons, black, 6.1% (SD 5.1, p = 0.23). Cell counts are from individual wells for four separate differentiations.

See also Figure S4. Error bars represent SEM.

potential back to negative values after an action potential, a decrease in such currents likely contributes to increased action potential firing in ALS motor neurons.

Correction of the disease-causing SOD1A4V/+ mutation also increased the relative steady-state delayed-rectifier potassium current amplitude, showing that this phenotypic difference specifically resulted from the A4V mutation (p < 0.005, t test; Figure 2F). We found a marked reduction in delayed-rectifier current magnitude in 39b compared to 39b-Cor motor neurons (p < 0.05, t test; Figure 2G) but no difference in sodium current peak amplitudes (p = 0.8, t test; Figure 2H). Thus, correction of the deficit in delayed-rectifier potassium current in 39b-Cor motor neurons may enable repolarization of the membrane potential back to normal hyperpolarized values and reduction of excitability to levels in wild-type motor neurons.

Retigabine Blocks Motor Neuron Hyperexcitability and Increases In Vitro Survival of SOD1A4V/+ ALS Motor Neurons

Motor neurons express many types of voltage-activated potassium channels and pharmacological dissection and quantification into distinct components is challenging. Regardless of which currents produce the hyperexcitability in diseased motor neurons, Kv7 (KCNQ) channels are attractive targets for correcting the hyperexcitability because of their activation at subthreshold voltages and subsequent powerful control of excitability (Brown and Passmore, 2009). Given this and the reduced delayed-recti-fier potassium currents in ALS-derived motor neurons, we hypothesized that retigabine, a specific activator of subthreshold Kv7 currents and clinically approved anticonvulsant (Porter et al., 2007), might block hyperexcitability in the SOD1A4V/+ motor neurons. In whole-cell patch clamp, retigabine significantly increased the minimal current step necessary for action potential generation (rheobase) by 3.6 ± 2.4 pA (p < 0.05, Wilcoxon signed ranktest; Figure 3A). Retigabine also stopped spontaneous firing of Hb9::RFP-positive motor neurons and hyperpolarized the resting membrane potential by 6.0 ± 2.2 mV (p = 0.001, t test; Figure 3B). Because these experiments were performed with blockers of glutamatergic, GABAergic, and glycinergic receptors, retigabine must have a direct effect on motor neuron excitability. We used MEA recordings to determine a dose-response for inhibition of spontaneous firing by retigabine of SOD1A4V/+ ALS-derived neurons. Retigabine suppressed ALS neuron spontaneous firing with an EC50of 1.5 mM (Figure 3C), a concentration consistent with its pharmacological activity as an antiepileptic agent and similar to its EC50 for Kv7 channels (Wickenden et al., 2000). In line with this finding, analysis of RNA sequencing (RNA-seq) data from FACS-sorted motor neurons (Kiskinis et al., 2014) confirm expression of Kv7 channels (Table S3).

To evaluate the possibility that hyperexcitability is an upstream modulator of motor neuron degeneration in ALS, we tested if retigabine affects the survival of control and SOD1A4V/+ motor neurons over 30 days in culture. As observed by Kiskinis et al. under basal conditions, the loss of SOD1A4V/+ motor neurons was greater than SOD1+/+ control motor neurons (Kiskinis et al., 2014). Two weeks of treatment with retigabine (1 mM) increased the number of ALS motor neurons in vitro by 25% (p < 10—4, t test; Figure 3D) to levels found in controls.

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Figure 4. Hyperexcitability of C9orf72 Repeat Expansion-Derived Motor Neurons

(A) Representative recordings from four/64 MEA electrodes recorded from control (11a, 18a) and C9orf72 expansion ALS-derived neurons (19f, RB8b) cultured for 14 days.

(B) Total action potential firing rate during 1 min of recording from MEAs (11a, n = 1; 18a, n = 3; control mean 4,752 ± 2,786 spikes/min; 19f, n = 3; RB8b, n = 3; ALS mean 20,022 ± 3,775 spikes/min; p < 0.05, t test).

(C) Average of mean firing rate for control and C9orf72-derived neurons (11a, n = 82; 18a, n = 203; control mean 1.11 ± 0.06 Hz; 19f, n = 407; RB9d, n = 929; ALS mean 1.50 ± 0.04; p < 10~5, t test).

Error bars represent SEM.

To investigate how retigabine increases the survival of SOD1A4V/+ ALS motor neurons, we determined whether it affects pathways suspected to contribute to motor neuron death in ALS (Robberecht and Philips, 2013). We chose to look at endoplasmic reticulum (ER) stress because of the demonstration that ER stress pathways are activated in SOD1A4V/+ ALS compared to SOD1+/+ motor neurons (Kiskinis et al., 2014). After 2 weeks of treatment with retigabine (1 mM), XBP1 splicing was markedly decreased in retigabine compared to vehicle-treated 39b SOD1A4V/+ ALS motor neurons (Figures S4A and S4B). In addition to reduced XBP1 splicing, we observed a decrease in PUMA and increase in EIF2B3 transcript levels, consistent with

downregulation of ER stress in response to retigabine treatment (Figure S4C).

Motor Neuron Hyperexcitability Is Present in Distinct ALS Forms and Is Blocked by Kv7 Activators

In order to investigate whether motor neuron hyperexcitability generalized to additional ALS variants, we performed MEA recordings of motor neurons derived from iPSC lines made from two unrelated familial ALS patients with C9orf72 hexanucleotide repeat expansion (19f and RB8b) (Kiskinis et al., 2014), which is responsible for 40%-50% of familial ALS and approximately 10% of sporadic cases (Robberecht and Philips, 2013). Motor neurons derived from these patients also showed significant hy-perexcitability compared to controls in both total firing rate (p < 0.05, t test; Figures 4A and 4B) and average mean neuronal firing rate (p < 10~5, t test; Figure 4C).

We reasoned that comparing neuronal firing properties of a large group of ALS patient and control-derived motor neurons would help evaluate the robustness of the motor neuron hyperexcitability and, together with the A4V gene correction experiments, eliminate artifacts due to cell line variation. SOD1-derived motor neurons (four lines from four unrelated subjects harboring three different mutations), C9orf72-derived motor neurons (two lines from two unrelated subjects), fused-in-sarcoma (FUS)-derived motor neurons (two lines from two unrelated subjects harboring two different mutations) were all hyperexcitable relative to motor neurons derived from six iPSC lines made from five individual healthy controls (ANOVA, p < 10~7; Tukey's post hoc tests for control versus SOD1 p < 0.01, control versus C9orf72 p < 0.01, control versus FUS p < 0.05; Figure 5A). Furthermore, spontaneous action potential firing in the ALS variant-derived motor neurons was uniformly blocked by retigabine (Figure 5B). Consistent with an on-target effect of retigabine, we found that a chemically distinct but less potent Kv7 current enhancer, flupirtine (Brown and Passmore, 2009), also blocked spontaneous motor neuron firing (Figure 5C). These results demonstrate the broad relevance of motor neuron hyperexcitability for familial ALS and its sensitivity to Kv7 agonists across iPSC lines, patients and genotypic etiologies.


Neurons derived from patient iPSCs can be used to investigate physiological changes in specific neural subtypes relevant to neurodegeneration and reveal important disease mechanisms and candidate therapeutics. We found consistent hyperexcit-ability in motor neurons from a broad group of familial ALS patients, whose disease-causing mutations collectively span the majority of familial ALS cases. Differential excitability of particular motor neurons has been proposed to explain the selective vulnerability of specific motor neuron pools in ALS, and our data provide a possible mechanistic basis for this hypothesis (Bae et al., 2009; Saxena and Caroni, 2011).

Motor neuron hyperexcitability may contribute, therefore, to motor neuron death, although connections between hyperexcitability and motor neuron death in other ALS variants will now require investigation. Our results are consistent with multiple

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studies of excitotoxicity (Cleveland and Rothstein, 2001; Fritz et al., 2013) but, potentially, not with two recent studies. One postulated that increased excitability in the SOD1G93A mouse model may be a compensatory mechanism (Saxena et al., 2013). However, systemically administered glutamatergic and cholinergic modulators may affect multiple neuronal types, making it difficult to assess how the findings relate to motor neuron excitability, which was not measured. A second study found decreased numbers of elicited spikes in iPSC-derived motor neurons from ALS patients with C9orf72 repeat expansion compared to controls (Sareen et al., 2013). The differences in spike count here may reflect the longer differentiation time (6679 days), differences in surviving neuronal populations or alterations in resting membrane potential in C9orf72 compared to control neurons. For example, the more depolarized resting potential in C9orf72 expansion-derived neurons could result in greater sodium channel inactivation and reduced capacity for generation of multiple action potentials; indeed, depolarization-induced action potential blockade may be a late phase of progressive hyperexcitability. Mouse SOD1G93A motor neurons show hyperexcitability even at an embryonic age (Kuo et al., 2004; Pieri et al., 2003; van Zundert et al., 2008), and neurophys-iological studies reveal increased excitability in C9orf72 repeat expansion subjects (Williams et al., 2013) in addition to other familial and sporadic ALS cases (Blair et al., 2010; Mills and Nithi, 1997; Vucic and Kiernan, 2006, 2010).

Figure 5. Motor Neuron Hyperexcitability and Block by Retigabine Are Broad Properties of ALS Variants

(A) Multielectrode array recordings of motor neurons derived from control (11a, n = 8; 15b, n = 2; 17a, n = 5; 18a, n = 7; 18b, n = 10; 20b, n = 6), SOD1 (25b, D90A, n = 3; 27d, G85S, n = 7; 39b, A4V, n = 3; RB9d, A4V, n = 7), C9orf72 expansion (19f, n = 2; RB8B, n = 13) and FUS (MGH5b, frameshift mutation at residue 511, n = 10; RB21, H517Q, n = 4) subjects cultured for 14 days. ANOVA, p < 10~7; Tukey's post hoc tests for control versus SOD1 p < 0.01, control versus C9orf72 p < 0.01, control versus FUS p < 0.05. For subject 18, motor neurons from two different iPSC lines were recorded. Error bars are 95% confidence interval. See also Figure S5.

(B) Dose-response curve for retigabine on suppression of spontaneous action potentials in MEA recording and Hill plot fit of mean data from SOD1 (n = 10; EC50 1.9 ± 0.5 mM), C9orf72 (n = 9; EC50 2.6 ± 0.8 mM) and FUS (n = 4; EC50 1.9 ± 1.1 mM).

(C) Dose-response curve for flupirtine on suppression of spontaneous action potentials in MEA recording and Hill plot fit of mean data from SOD1 (n = 5; EC50 9.8 mM), C9orf72 (n = 4; EC50 19.4 mM) and FUS (n = 2; EC50 15 mM).

Error bars for (B) and (C) represent SEM.

Previously, attention has focused on Flupirtine (pM) persistent sodium currents as a mecha-

nism of motor neuron hyperexcitability (Kuo et al., 2005; Vucic and Kiernan,

2010), and riluzole, the only approved drug for ALS, blocks this current (Urbani and Belluzzi, 2000). Ourfindings suggest an additional important role of voltage-activated potassium channels. Channel compartmentalization may differ between in vivo and cultured-neuron systems; however, a recent immunohistochem-ical study of human spinal cords found decreased protein levels of a delayed-rectifier potassium channel selectively in the ventral roots of sporadic ALS but not control subjects (Shibuya et al.,

2011). Mutations in voltage-gated potassium channels cause other neurodegenerative diseases (Waters et al., 2006), and oxidation of potassium channels (with consequent modulation of voltage-dependence and kinetics of channel gating) has been suggested as a broad mechanism of aging and neurodegeneration (Sesti et al., 2010). DPP6, which is involved in trafficking Kv4 voltage-gated potassium channels in CA1 hippo-campal neurons, is linked to ALS (Sun et al., 2011; van Es et al., 2008). Hypermethylation and downregulation of potassium channel genes were observed in epigenetic analyses of postmortem ALS patient spinal cords (Figueroa-Romero et al., 2012).

Both enhanced persistent sodium and reduced potassium currents could converge to produce hyperexcitability, and both may offer complementary pharmacological targets to control it. Although our study evaluated hyperexcitability as an innate or autonomous property of motor neurons, differences in excitability may also reflect interactions between motor neurons and glia or Schwann cells (Fritz et al., 2013). Hyperexcitability

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has also been observed in motor neurons from a mouse model of spinal muscular atrophy (Mentis et al., 2011), and there is evidence for both cell autonomous (Gogliotti et al., 2012) and nonautonomous (Imlach et al., 2012) modulation of excitability. Interestingly, spinal muscular atrophy has also been linked to DPP6, raising the possibility of potassium channel contributions (van Es et al., 2009).

The pathways connecting disease-causing mutations and decreased potassium channel function, and between hyperexcitability and motor neuron death remain to be clarified. Calcium overload through voltage-gated calcium channels (Chan et al., 2007) and activation of ER stress (Saxena and Caroni, 2011) are possibilities. The finding that decreasing motor neuron activity reduces ER stress suggests that hyperexcitability may be upstream of the unfolded protein response, explaining at least partially how hyperexcitability may contribute to motor neuron death in ALS. Because ER stress modifiers increase motor neuron activity, a vicious cycle may result in ALS from reciprocal positive feedback between hyperactivity and ER stress (Kiskinis etal., 2014).

Despite the asymptomatic early decades typical of ALS patients, we observed a disease-specific phenotype in iPSC-derived neurons cultured for only weeks. That this property manifests so quickly in culture may reflect the absence of supporting cells and the inhibitory circuitry normally present in vivo. Identification of a screenable electrophysiological phenotype that contributes to motor neuron death and manifests quickly could facilitate investigation of ALS pathophysiology and identification or validation of therapeutics for individual patients. Furthermore, the phenotype may offer a personalized medicine approach to treatment, using response of stem cell-derived neurons as a guide for individual patient treatment selection; this strategy will require motor neurons derived from large cohorts of patients for validation. Studies of motor neuron excitability can now be expanded to determine whether all forms of ALS converge onto a single mechanistic pathway. More generally, our study illustrates the potential for using iPSCs differentiated into specific disease-relevant cell types to identify disease phenotypes, novel biomarkers, and potential treatments.


iPSC Lines, Culture, and Motor Neuron Differentiation

iPSC generation from patient fibroblasts obtained under institutional review board approval, characterization, and motor neuron differentiation were performed as described in Kiskinis et al. (2014) and Figure S5, with iPSC line scoring as done previously (Bock et al., 2011). For transfection of 18a and 39b lines with a Hb9::GFP reporter (Figure S3), a 1 kb Hb9 promoter fragment (gift from Hynek Wichterle) controlling the expression of myristoylated GFP was inserted into a donor plasmid specific for the AAVS1 locus (Sigma). Subsequently, 2.5 million iPS cells were accutased and electroporated using the Neon transfection system (100 ml tip; 1,600 V voltage, 20 ms width, 1 pulse; Life Technologies) with 2 mg of AAVS1 ZFN plasmid and 6 mg of modified AAVS1 donor plasmid. After nucleofection, cells were plated on matrigel with mTeSR1 in the presence of ROCK inhibitor. After 48 hr, puromycin selection was applied, and surviving clonal colonies were individually passaged and genomic DNA was extracted. PCR was used to confirm proper targeting of the cassette. Primer sequences are available upon request. Faithful expression of the reporter was verified using expression of the motor neuron marker Isl1 (Figure S3C).

iPSCs were maintained on culture dishes as described previously (Boulting et al., 2011) with modifications (Kiskinis et al., 2014) in a 24 day protocol based on initial neuralization with SB431542 (10 mM, Sigma-Aldrich) and Dorsomor-phin (1 mM, Stemgent), and motor neuron patterning with RA (Sigma) and a small smoothened Agonist 1.3 (Calbiochem). For Figure 1F, FACS-purified neurons were grown on a confluent monolayer of primary cortical mouse glia prepared from P0-P2 mouse pups (as described in Boulting et al., 2011), which may increase firing rates compared to experiments without glia (Boehler et al., 2007).

MEA Recording

After 24 days of differentiation, equal numbers of control and ALS neurons were plated on poly-D-lysine/laminin coated p515A probes (Alpha Med Scientific) or M768-GLx 12-well plates (Axion BioSystems) at typical densities of 40,000-80,000/ probe or well. All probes were visualized immediately before each recording session to confirm a full monolayer of cells. Initial experiments (11a, 18a, 39b, and RB9d comparison) were performed as close as possible to the time of patch recordings (4 weeks). However, because we wished to evaluate firing at a time point prior to significant motor neuron death (Kiskinis et al., 2014), we performed subsequent experiments (39b-Cor and 39b comparison and all later experiments) at day 14 after dissociation (Table S1).

Recordings from 64 extracellular electrodes were made using a Med64 (Alpha Med Scientific) or Maestro (Axion BioSystems) MEA recording amplifier with a head stage that maintained a temperature of 37°C. For Med64 recordings, data were sampled at 20 kHz, digitized, and analyzed by spike clustering and spike extraction algorithms using Mobius software (Alpha Med Scientific) with a 2 kHz 9-pole Bessel low pass filter, 10 mV action threshold detection limit, and 30% cluster similarity radius. These standard settings were maintained for all analyses. We confirmed that we obtained similar results across a wide range of action potential threshold and cluster similarity radius settings. Correlation analysis to detect and correct for clusters detected by multiple electrodes was performed using custom MATLAB software. Total action potential firing rates and mean neuronal firing frequencies were then determined and plotted. In orderto record in larger replicates, we used the Axion Maestro MEA device, in a 12-well format with 64 extracellular electrodes in each well. For Maestro recordings, data were sampled at 12.5 kHz, digitized, and analyzed using Axion Integrated Studio software (Axion BioSystems) with a 200 Hz high pass and 2.5 kHz low pass filter and an adaptive spike detection threshold set at 5.5 times the SD for each electrode with 1 s binning. These standard settings were maintained for all Axion MEA recording and analysis.

For retigabine dose-response curves, action potential numbers during 1 min of recording in each concentration of retigabine were normalized to the initial action potential number during 1 min of recording in standard extracellular saline solution. The EC50 value was determined by fitting the mean normalized data values to the Hill equation, y = 1/((EC50/x) nH+1) where nH is the Hill coefficient.

Patch Electrophysiology

Whole-cell patch recordings were performed on iPS-derived motor neurons identified by transduction with an Hb9::RFP lentivirus. Lentiviral transduction was typically performed 7-10 days before recording. Two large comparisons were performed, one consisting of 11a, 18a, 39b, and RB9d, and the second consisting of 39b-Cor and 39b. Each comparison was made from pooled data from multiple separate experiments, each consisting of synchronous and parallel iPSC culture and differentiation, embryoid body dissociation, plating, and maturation of control-and ALS-derived neurons. Equal numbers of control and ALS motor neurons were recorded from each experiment. Comparison of the original four lines (11a, 18a, 39b, and RB9d) was made using four separate parallel differentiation experiments, whereas comparison of the isogenic correction comparison (39b-Cor and 39b) was performed using three separate parallel differentiation experiments.

For each experiment, neurons from control and ALS lines were dissociated after 24 days of differentiation and plated onto poly-d-lysine/laminin coated glass coverslips (20,000-40,000/coverslip) and allowed to mature for 4 weeks from start of differentiation. We chose 4 weeks as the best time point because this yielded the most homogeneous population of mature-appearing Hb9::RFP-positive motor neurons (at the requisite low cell density for patch

clamp) with the most mature electrophysiological properties. Whole-cell current-clamp and voltage-clamp recordings were performed using a Multiclamp 700B (Molecular Devices) at room temperature (21°C-23°C). Data were sampled at 20 kHz and digitized with a Digidata 1440A A/D interface and recorded using pCLAMP 10 software (Molecular Devices). Data were low-pass filtered at 2 kHz. Patch pipettes were pulled from borosilicate glass capillaries on a Sutter Instruments P-97 puller and had resistances of 2-4 MQ. The pipette capacitance was reduced by wrapping the shank with Parafilm and compensated for using the amplifier circuitry. Series resistance was typically 5-10 MQ, always less than 15 MQ, and compensated by at least 80%. Neurons were excluded from analysis if holding current at —80 mV exceeded 100 pA, input resistance was less than 250 or greater than 2000 MQ, or spikes elicited from —65 mV had peaks below 0 mV. Resting membrane potential was determined by averaging for 20 s of recording, and afterward a small holding current (typically with amplitude less than 5 pA) was used to clamp the resting membrane potential as close as possible to —65 mV. Rheobase was measured by applying 1 s steps in increments of 2.5 pA until an action potential was generated. Current ramps were elicited from an initial hyperpolarizing current of 10 pA for 1 s followed by a 210 pA/s depolarizing ramp of duration 1 s. Spikes on the ramps were counted if the peak voltage exceeded —10 mV. Action potential properties (Table S2) were determined using custom-written analysis software in Igor Pro (Wavemetrics) with DataAccess (Bruxton) for importing the files. For voltage-clamp recordings, voltages were elicited by 100 ms depolarizing steps from a holding potential of —80 mV to test potentials ranging from —80 mV to 50 mV in 10 mV increments. For the latter gene correction experiments, the step length was increased to 200 ms to assay delayed-rectifier currents after more complete decay of transient potassium currents. For retigabine patch applications, resting membrane potential was recorded immediately before and 10 s after the application of 10 mM retigabine (in each case, membrane potential was an average of values sampled for 20 s). For all patch experiments, series resistance was monitored by brief —5 pA hyperpolarizing steps during current clamp recordings and by 5 mV hyperpolarizing steps during voltage-clamp recordings. Electrode drift was measured at the end of each recording and was typically 1-2 mV. The extracellular solution was sodium-based and contained 135 mM NaCl, 5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 10 mM glucose, 10 mM HEPES 10 (pH 7.4). The intracellular solution was potassium based and contained 150 mM KCl, 2 mM MgCl2,10 mM HEPES, 4 mM MgATP, 0.3 mM NaGTP, 10 mM Na2PhosCr, 1 mM EGTA (pH 7.4). For isolation of delayed-rectifier potassium channels (Figure 2G), 300 nM TTX and 100 mM CdCl2 were used to block voltage-gated sodium channels and voltage- and calcium-activated potassium channels, respectively. For isolation of voltage-gated sodium currents (Figure 2H), internal KCl was replaced by CsCl to block potassium currents and 100 mM CdCl2 was used to block calcium currents.

Cell Survival and ER Stress Assays

Dissociated neurons (20,000) were plated on poly-D-lysine/laminin coated 8-well chamber slides (BD Biosciences) containing a confluent monolayer of primary cortical mouse glia (Boulting et al., 2011). For survival analysis, slides were fixed at 3 and 30 days, and cultures were stained for counting. The number of ISL-positive, TUJ1-positive motor neurons (counter blinded to cell line identity) was normalized to the number on day 3 for each line. Retigabine (1 mM) or vehicle control was added from day 15 onward. Eleven total experiments (control motor neurons) and nine total experiments (ALS motor neurons) were performed from the same four separate differentiations. ER stress experiments were performed as described in Kiskinis et al. (2014). In brief, in two separate independent biological replicates, 300 ng of RNA was used to generate cDNA, of which 2 ml and AmpliTaq Gold Polymerase (Applied Biosystems) were used for PCR amplification. The amounts of spliced and unspliced bands were quantified using Image J. Quantitative RT-PCR was performed intriplicate using the iSCRIPT kit (Bio-Rad) for cDNA synthesis and SYBR green (Bio-Rad) labeling followed by amplification using the iCycler system (Bio-Rad).

Drugs included retigabine (Santa Cruz Biotechnology), bicuculline, strychnine, D-AP5, CNQX, flupirtine, TTX (all from Tocris Bioscience). CsCl and CdCl2 were from Sigma.

Statistical Analysis

p < 0.05 was considered statistically significant. Comparisons were made between control and ALS populations using t tests (two-tailed, unpaired)/ANOVA for continuous data and rank tests for nonparametric data (discrete measurements of number of spikes on ramps and rheobase). For analysis of MEA firing over time (Figure 1F), we used a mixed repeated-measures ANOVA model with fixed effects of cell line and time and random effects of individual replicate. For effect of retigabine on specific cells (Figures 3A and 3B), paired tests were used. For effect of retigabine on survival (Figure 3D), we fitted a linear regression model with effects of retigabine treatment, cell line, and their interaction. F-tests for difference between ALS and control lines gave p = 0.0011, for effect of retigabine p = 3.8 x 10~4, and for effect modification p = 0.015. For ALS subjects, the effect of retigabine was an increase in cell count of 25.3% (SD 5.6, p = 6.4 x 10~5). The effect estimate for 39b was 16.9% (SD 12.9, p = 0.03) and for RB9d 39.9% (SD 11.3, p = 0.001). For control subjects, the effect of retigabine was an increase in cell count of 6.1% (SD 5.1, p = 0.23). For multiple genotype comparisons (Figure 5A), one-way ANOVA after log transformation to normalize variance and post hoc Tukey tests were used to analyze multiple ALS variants. Error bars represent SEM unless indicated.


Supplemental Information includes five figures and three tables and can be found with this article online at


We thank W. David and S. Cash for comments, suggestions, and review of the manuscript, K. Kapur for assistance with statistical analysis, and K. Wainger for assistance with figures. This work was supported by NIH 5 T32 GM007592-33, Harvard NeuroDiscovery, ALS Association, and American Brain Foundation Clinical Research Fellowship (B.J.W.); Charles King Trust Postdoctoral Fellowship (E.K.); American Brain Foundation/ALS Association and KL2 MeRIT fellowship/Harvard Catalyst (S.S.W.H.); ALS Therapy Alliance, P2ALS, Angel Fund, Pierre L. de Bourgknecht ALS Research Foundation, Al-Athel ALS Research Foundation, ALS Family Charitable Foundation and NIH/NINDS (1R01NS050557 and NINDS ARRA Award RC2-NS070-342) (R.H.B.); P2ALS, Project ALS, Target ALS, NINDS GO grant (5RC2NS069395-02), NINDS R24 (1U24NS078736-01) and HHMI (K.E.); NIH (5 R01 NS038253-10; 2 R01 NS038153-15), Target ALS and New York Stem Cell Foundation (C.J.W.). GlaxoSmithKline supports research distinct from this study as part of an alliance with the Harvard Stem Cell Institute, including the Woolf and Eggan labs.

Received: September 30, 2013 Revised: February 17, 2014 Accepted: March 10, 2014 Published: April 3, 2014


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